QGRS-mapper identifies potential GQ-forming sequences in all RV genomes.
Employing extensive bioinformatics analysis, Lavezzo and colleagues recently predicted GQs in viral genomes of human virus species, including RVs 34. We independently confirmed and extended their findings for all RV genome sequences available in Genebank by using a local copy of the same program that runs on the quadruplex-forming G-rich sequence (QGRS) mapper web-based server 35. This software has been originally developed to identify putative QGRS in DNA, where the G-score reflects the stability of a predicted GQ, which usually increases with the number of G-tetrades and decreases with the loop size. The respective search motif G≥ 3L1−7G≥ 3L1 − 7G≥ 3L1 − 7G≥ 3 takes into account that most experimentally identified DNA GQs conformed to short-looped (1 to 7 nucleotides) structures with three and occasionally more G-tetrades. However, RNA GQs are generally more stable than the corresponding DNA GQs 36; two-layer RNA GQs are therefore not uncommon 37, and the overall higher stability of RNA GQs allows the insertion of larger loops compared to DNA GQs 38, 39. Recently, an atypical RNA GQ with no first loop has been described 37. Therefore, we adjusted the search parameters of the QGRS mapper to allow the identification of putative unconventional long-loop and zero-nucleotide loop G-quartets, including those featuring only two layers. We then plotted the respective QGRS prediction G-scores ≥ 10 against their positions in the genomic RNA sequences (Fig. 1a). Strikingly, the vast majority of rhinoviral genomes feature only QGRS predicted to form just two-layered GQs. RV-A41 and RV-B4 are singled out by the additional presence of one and two three-layer GQs, respectively.
Furthermore, most RV-ABC genomes feature at least one and up to seven putative zero-nucleotide loop GQs without marked conservation. The number of potential QGRS varies from 6 to 19 (mean 12) for the RV-As, from 9 to 19 (mean 13) for the RV-Bs, and from 10 to 23 (mean 15) for the RV-Cs. The augmented prevalence of QGRS for the RV-C species is likely attributable to their genomes' distinctly higher GC content (43 % for RV-C, 38 % for RV-A, and 38 % for RV-B 40). The analysis further revealed four highly conserved QGRS (Fig. 1a - asterisk), all located in the open reading frame. The slight differences in their position between the genera (RV-A, RV-B, and RV-C) result from clade-specific insertions and deletions. An additional conserved QGRS is uniquely present in the RV-Bs, upstream and close to the second highly conserved QGRS motif. By contrast, only moderately conserved QGRS were predicted in the 5´ untranslated region (UTR), which comprises essential regulatory elements such as the 5´ cloverleaf structure and the IRES, required for replication and cap-independent translation 41. No single QGRS was predicted within the short 3´ UTR, featuring a highly conserved hairpin involved in virus replication 42.
NMR analysis demonstrates the folding of synthetic ribooligonucleotides representing selected QGRS of RV-A2 RNA into GQs, which is differentially affected by PDS.
For RV-A2, QGRS-mapper predicted 11 QGRS with G-score ≥ 10 (Supplementary Fig. S1). We chose to study the candidates with the lowest and highest G-scores, G11 and G20, the latter also representing one of the four highly conserved GQs (see above). Assuming the almost invariable adoption of parallel GQs by RNA, G11 would give rise to an unusual monomeric two-layer GQ bearing a zero-nucleotide loop 3 in combination with a long loop 1 (Table 1; a schematic illustration of the likely G11 and G20 RNA GQ structure is displayed in Supplementary Fig. S1). To confirm their GQ-forming propensity, we collected 1H NMR spectra using 250 µM of the derived synthetic ribooligonucleotides in 10 mM sodium phosphate (pH 7.4), 100 mM KCl, similarly as described elsewhere 43. Each ribonucleotide was denatured at 95°C for 10 min, followed by cooling to 4°C for 10 min to favour intramolecular annealing into GQ and possibly other secondary structure(s) in the presence of monovalent cations. A high concentration of K+ usually stabilises GQs, as shown in other instances 36. G11 and G20 displayed 1H signals of the bulk of the imino proton in the region between 10.4–12.4 ppm, which is the typical NMR signature of GQs (Fig. 1b – salmon-coloured area). The lack of sharp signals can be explained by G-register exchange dynamics 44 since certain G runs have more than two Gs, indicating the possible presence of alternative conformations.
Peaks in the canonical Watson-Crick base pair region at 1H NMR shifts larger ~ 12.4 ppm (olive-coloured area) are observable at low (4°C) but not at high temperatures (34°C, the optimal temperature for RV-A2 replication), indicating that these secondary structure elements (presumably hairpin(s)) are less stable than the GQs. Within the range assigned to GQs, the second peak (11.9–12.4 ppm) observed only for G11 (Fig. 1b) exhibited unaltered intensity in an HDX experiment performed for 7 h at 37°C. The solvent inaccessibility of these imino protons indicated that they were most likely also part of the GQ core (Supplementary Fig. S2a). In the G20 spectrum, three distinct peaks (one within and two next to the upfield broader of the GQ region) completely disappeared with increasing temperature, indicating that the corresponding imino protons are conceivably not involved in the G-tetrad core formation. The addition of 500 µM pyridostatin (PDS), which specifically binds with high affinity to DNA and RNA GQs but not to other nucleic acid secondary structures except i-motif forming DNA 45, resulted in substantial changes of the imino 1H peaks for both synthetic RNAs (Fig. 1b and Supplementary Fig. S2b), indicative of a specific interaction.
The only other 1H-NMR study we are aware of using PDS and a GQ was performed with a three-layer DNA GQ derived from the src kinase gene (SRC). This showed an upfield-shifting of GQ imino proton signals stemming from PDS binding to the top G-quartet in a stacking mode 46. However, the differential effect observed by us suggested that the binding mode of PDS may vary for G11 vs G20. We hence used molecular docking to assess this possibility further. For lack of detailed structural information about the G11/G20 system, we instead in silico evaluated the interaction of a crystallographic model of a pseudorabies virus RNA GQ molecule composed of two G-quartets (PQS18-1, GGCUCGGCGGCGGA) (PDB 6JJH 47) and PDS. Using increasing concentrations of PDS in this in silico analysis, we unexpectedly discovered that it could form defined dimers mostly stabilised by π-π interactions. This was then experimentally substantiated by UV-absorption spectroscopy, which clearly demonstrated a concentration-dependent self-association of PDS (Ka ~42 µM) in the buffer chosen for NMR spectroscopy (Supplementary Fig. S3).
The analysis of the best docking poses for the monomeric and dimeric PDS and PQS18-1 revealed two distinct classes of binding modes (Supplementary Fig. S4): i) end-stack (a and c), and ii) groove/loop-binding (b and d). In both instances, the 2-(quinolin-4-oxy)ethanamine moiety of PDS interacts with the nucleobases via Van der Waals forces and an additional contribution of electrostatic interactions between the charged amino groups of PDS structure and the phosphate groups of the GQs. These binding modes may be variably exploited for the interaction with G11 and G20, presumably resulting in the idiosyncratic PDS-induced peak shifts.
Altogether, these results provided the initial bioinformatics analysis demonstrating the likely formation of two-layered GQs by both ribooligonucleotides under favourable conditions for the folding of such structures.
Table 1
– Sequences of the synthetic ribooligonucleotides used in this study. Note that in the negative control, all Gs of miniTERRA were replaced by Cs.
Ribooligonucleotides
|
Sequence
|
Negative Control
|
5‘- UUA CCC UUA CCC UUA CCC UUA CCC UUA − 3‘
|
miniTERRA (G-score = 42)
|
5‘- UUA GGG UUA GGG UUA GGG UUA GGG UUA − 3‘
|
G11 (position 2048–2074)
|
5‘- GGC ACU CAU GUU AUA UGG GAU GUG GGG − 3‘
|
G20 (position 1038–1064)
|
5‘- CCU CAA AGG GUU GGU GGU GGA AAC UAC − 3‘
|
Circular dichroism spectroscopy of the synthetic ribooligonucleotides G11 and G20 reveals a parallel GQ folding topology.
We next verified the tentatively assigned folding topology of G11 and G20 by circular dichroism (CD) spectroscopy. CD is a gold standard to determine the strand orientations of GQs. These might be parallel, as commonly adopted by RNA and DNA, or anti-parallel and hybrid, which, with a few exceptions 48, are observed only for DNA. Telomeric-repeat-containing RNA (miniTERRA; Table 1), which adopts a three-layer parallel conformation 49, served as a positive control.
The CD profiles of miniTERRA, G11, and G20 taken at 25°C in either 100 mM sodium or 100 mM potassium phosphate buffer (pH 7.4) in each instance displayed the signature of an all-parallel topology (ellipticity shows a negative band at 240 nm and a positive band at 265 nm). The monovalent Na+ and K+ cations differentially stabilise GQs owing to their different ionic radius and hydration free energies 50. Generally, potassium promotes folding and stabilises (ribo)oligonucleotide GQs to a larger extent than sodium 36, 51, though some exceptions have been reported 52. In accordance with the latter, the degree of GQ formation of G20 was noticeably the same irrespective of the presence of Na+ or K+ as indicated by the practically unchanged CD spectrum (Fig. 1c). By contrast, as found for the vast majority of GQ forming RNAs, the miniTERRA control and the G11 ribooligonucleotide showed a higher proportion of GQ structure in K+ compared to Na+. The trough at 210 nm only evident for G11 and G20 is likely attributable to the additional presence of structural elements comprising A-type duplex RNA regions as also revealed in the above NMR analysis at low temperature.
Na + and K+ differently impact the physicochemical parameters of G11, G20, and miniTERRA ribooligonucleotides.
As seen in Fig. 2a, the CD melting and annealing spectra are almost superimposable for the G11 and G20 ribooligonucleotides and the miniTERRA control when dissolved in a sodium-containing buffer. The lack of a marked hysteresis indicates thermodynamic equilibrium between denaturation and refolding in the examined temperature range and the used ramp rate and is typical for intramolecular GQs 53. The situation for miniTERRA was strikingly different in the potassium buffer, where the unfolding/annealing profiles exhibited a marked hysteresis (Fig. 2a); this is typically observed for intermolecular GQ formation, where the shape of the melting and refolding curves depends on the speed of temperature ramping 53. Another possibility is a slow conformational change of an intramolecular GQ from one topology to another 54. However, according to CD spectroscopy. miniTERRA in the presence of either cation (Na+, K+) formed a parallel GQ (Fig. 1c) exclusively. Since the multimerisation of miniTERRA could be ruled out by a subsequent electrophoretic analysis (see below), the relatively slow ramp rate of 1°C/min apparently still did not allow the reversible refolding of this three-layer intramolecular GQ. This is in line with findings by others that multiple layers of tetrades are particularly prone to hysteresis 55. By contrast, in the presence of K+, the CD melting/cooling profiles of both two-layered GQs exhibited just a rather subtle (G11) to negligible hysteresis (G20). The latter's overlapping heating and cooling curves, as also seen with Na+ buffer, reinforces the notion that G20 forms a monomeric (see below) intramolecular GQ in presence of each of these cations.
Others have shown that K+-containing solutions stabilise the parallel topologies of two- and three-quartet RNA GQ much more than Na+-containing solutions, with the difference in the melting temperatures ΔTm ranging from 15 to more than 30°C 36, 51. Such drastic change in Tm (determined at the intersection of the second derivative with the x-axis in Fig. 2a) was also evident for the three-layer miniTERRA GQ in K+ compared to Na+. By contrast, the melting temperatures of G11 and G20 in the K+ containing buffer were just ~ 4°C higher than those determined in Na+ buffer.
The limited hysteresis of G11 melting/cooling might indicate the formation of a dimeric GQ resulting from end-to-end stacking of two intramolecular GQs. This gains additional solid support from native gel electrophoresis, which clearly shows the occurrence of dimers in the presence of potassium ions for G11. Under these conditions, G20 and the miniTERRA control gave rise to just a single band, indicating that they existed primarily as monomers (Supplementary Fig. S5). Based on these results, the slight hysteresis of the G11 melting can be best interpreted by assuming a fast monomer refolding, followed by slower oligomerisation.
The adoption of a GQ conformation by the two RV-A2 derived RNA sequences (G11 and G20) and the positive control (miniTERRA) was then confirmed with a Thioflavin T (ThT) light-up assay with all measurements done at room temperature. ThT end-stacks to RNA GQs, which strongly increases its fluorescence 56. As seen in Fig. 2b and 2c, no significant fluorescence was detected upon incubation with the negative control (C for G substituted miniTERRA) ribooligonucleotide (see Table 1). In contrast, a clear signal was obtained for all three tested GQs, whose magnitude was quite comparable for G11 in Na+ (left panel of Fig. 2b) and K+ buffer and about 1.5 and 2-fold higher in the presence of K+ for miniTERRA and G20, respectively (left panel of Fig. 2c). The ThT fluorescence intensity was strongest for miniTERRA and approximately 3 to 4-fold lower for G11 and G20 (but still 30 to 40 times higher than for the negative control), roughly correlating with their respective G-scores. Since the same amounts of oligonucleotides were employed, this indicated that a fraction of G11 and G20 adopted a non-GQ conformation as already observed in the 1H-NMR analysis. The coexistence of A-type RNA (hairpin) structure and a two-layer GQ structure has been recently described by Lightfoot and others 37. Based on the ThT light-up probe, G11, but not G20 would be almost indifferent to the choice of the two cations, which is opposite to the results of the label-free CD analysis. Most likely, the conformation of the respective GQs subtly differs when bound to Na+ (which coordinates with the Gs in the middle of a tetrade) or K+ (residing between G-tetrads), which may variably affect the binding affinity for ThT giving e.g. the impression of an apparent higher stability of G20 in K+ buffer. However, this does not change our overall conclusion that GQ structures are formed with both cations in this assay.
We then examined whether the tetrade-associated cation type (K+ or Na+) impacts the interaction of the respective GQs with PDS by conducting a fluorescent indicator displacement (FiD) assay, similar to the one described in refs. 57, 58. FID allows evaluating the relative affinity and selectivity of compounds binding to a GQ 59. Expectedly, ThT was displaced by PDS in each instance (right panels in Fig. 2b/c). However, while the dose-response curve for miniTERRA was comparable in Na+ and K+ containing buffers, the reduction in fluorescence determined for G11 and G20 was about 2-fold more efficient in the presence of Na+ as indicated by the respective IC50 values (Table 2). Notably, with the lower G-scoring G11, a sharp decrease of the ThT fluorescent signal was already evident at low PDS concentrations. Since ThT was shown not to markedly alter the stability of GQs 60, this might indicate a higher affinity of PDS for the noncanonical GQ-forming G11 in comparison to miniTERRA and G20 as reflected by the correspondingly lowest IC50 values (Table 2).
Table 2
– IC50 values for the displacement of ThT by PDS. Significance determined by using ANOVA with Tukey's multiple comparison test.
Ribooligonucleotides
|
IC50 (µM)
Na+ K+ Significance
|
mini TERRA
|
19.3 ± 0.5 17.5 ± 0.7 NS
|
G11 (position 2048–2074)
|
2.8 ± 1.4 6.1 ± 1.1 p < 0.0001
|
G20 (position 1038–1064)
|
10.5 ± 5.2 20.3 ± 1.0 p < 0.001
|
Pyridostatin (PDS) and PhenDC3 reduce RV-A2 infectivity.
Taken together, employing several orthogonal assays, we could demonstrate the intrinsic ability of two selected RV-A2-derived QGRS to form RNA GQs. However, within the context of the viral RNA, these and the other predicted candidate sequences may fail to fold into such scaffolds if embedded into or overlapping with alternative secondary structures of higher stability or more rapid formation (and trapped in a metastable state). Furthermore, the G-quartet fold of some untested putative QGRS may be too unstable at the temperature of virus propagation. PDS and other so-called GQ-stabilising compounds can selectively enhance the mechanical and thermal stability of DNA and RNA GQs over the level achieved by K+ alone 61, 62. Thereby, they can also force alternative secondary structures to transform into the compound-stabilised quadruplex conformation 63. The latter was evident for G20 and, less pronounced, for G11 by the loss of Watson-Crick peaks in the 1H-NMR spectrum on incubation with PDS (Supplementary Fig. S2b).
It is presently believed that the uncoating of RV-A2 and several other enteroviruses requires a structural switch of the genomic RNA and the transient unfolding of secondary structures for transit as a single-strand through one of the narrow pores formed in the A-particles 5, 7, 64, 65, 66. Therefore, we reasoned that the exposure of the encapsidated viral RNA to PDS by stabilising preexisting GQs and promoting the transition of unstructured and/or alternatively folded QGRS into such unconventional secondary structures might interfere with its in vitro and in vivo uncoating. RVs, in contrast to other enteroviruses such as poliovirus, are readily permeable for monovalent cations such as Cs+ already in the cold and also for small organic compounds such as dansylaziridine and ribogreen when incubated at breathing conditions 67, 68, 69. Capsid breathing describes a transitory expansion of the protein shell with temporary exposure of normally internal amino acid sequences through reversibly formed small holes, commencing at room temperature to around physiological temperatures, dependent on the RV serotype 70. We thus attempted to deliver PDS to the viral RNA genome within the native virion by exploiting this phenomenon. All incubations were done in phosphate-buffered saline (PBS) to prevent PDS aggregation into long fibrils, as we have recently observed for Tris- but not phosphate-based buffers 71.
First, we confirmed the dependence of PDS delivery to the inside of the capsid as a function of temperature. Purified RV-A2 was incubated with PDS under conditions of strongly diminished capsid breathing (at 4°C) and a capsid breathing-promoting temperature (at 34°C). Unbound PDS was removed, and a Particle Stability Thermal Release Assay (PaSTRy) 72, 73 was performed to determine a possible impact of PDS on temperature-dependent uncoating, a commonly used model for in vitro uncoating of picornaviruses 74. Figure 3a shows the temperatures where the genomic RNA becomes accessible to SYTO 82 as deduced from the temperature vs fluorescent emission curves 73 (Supplementary Fig. S6). Virus pre-incubated with PDS at 34°C exhibited a striking 4.7°C earlier onset of genomic RNA accessibility to SYTO 82 compared to the control condition, i.e., incubation with PDS at 4°C (to prevent diffusion of this compound through the protein shell) (Ton of 37.7°C vs. 42.4°C). However, the temperatures Tmax at the peak of the SYTO 82 signal (indicative of the complete conversion of all native virions into permanently porous A-particles) and the peak of the first derivative (at T50, corresponding to 50 % RNA accessibility) 73 remained practically unaltered.
We interpret this finding as that PDS, by interacting with (potential) QGRS in the genome of RV-A2, directly or indirectly affected RNA contacts with the capsid, thereby resulting in enhanced mobility of the protein shell of the native virus. This allowed appreciable uptake of the SYTO 82 dye into the viron for binding the viral RNA already at a lower temperature compared to the control. However, the PDS-induced effect did not critically impact the rate of the temperature-dependent conversion of native to A-particle as inferred from the unchanged T50 for this sigmoidal conversion 73. The heat-triggered RNA release starting at Tmax (resulting in the subsequent drop of the SYTO 82 signal) was also not affected. As thermal unzipping of secondary structures is required for the exit of the RNA from the capsid in this in vitro uncoating model 64, the bound PDS evidently did not increase the melting temperature of the viral GQs above one of the most stable non-GQ secondary structures (presumably mostly hairpin loops 75) inside the capsid.
We then assessed the effect of PDS on the in vivo uncoating of RV. HeLa cells were infected with RV-A2 pre-incubated with PDS and without PDS (control), both at 34°C for 4 h. This treatment time sufficed for the entry of appreciable amounts of RiboGreen into the capsid of RV-A2 69. Unbound PDS was removed by centrifugation in an Amicon ultrafilter unit, followed by repeated washing with PBS at 25°C. The viral samples were then transferred to HeLa cells grown in 10 cm diameter dishes and incubated for 30 min to allow for viral uptake and uncoating in the absence of inhibitory compounds 65. Supernatant and cells were collected by scraping, and internalised viral material was recovered by five times freezing and thawing. Cell debris was removed, and aliquots of the supernatants were subjected to immunoprecipitation with the subviral (A- and B-particle) specific monoclonal antibody 2G2 76, 77. From equal aliquots of the precipitated material, protein and RNA were recovered and quantified in Western blots (Fig. 3b) and by RT-qPCR, respectively (Fig. 3c). Comparing these results, it becomes clear that regardless of whether the pre-incubation was carried out in the presence or absence of PDS, the same amount of viral proteins were detected. This excludes any influence of PDS on cell attachment, e.g. via forming virus-trapping filaments as observed in Tris-based buffers (see above and ref. 71) or on the overall rate of native to subviral (A + B) particle conversion. However, quantification of the viral RNA contained in the 2G2 precipitates showed that the virus incubated in the presence of PDS and recovered from cells 30 min post-infection (pi) in the form of subviral A- and B-particles contained about 70 % more viral RNA than virus pre-incubated without PDS. This can be taken to indicate that significantly less RNA was released from the endocytosed PDS-treated virus during the first 30 min following the challenge of the cells.
Thirty min pi aliquots from similarly infected cells (as above) were also analysed by sedimentation through preformed sucrose density gradients. Ultracentrifugation under the specified conditions allows for separating native virus, A-particles, and empty B-particles 78. The profile in Fig. 3d verifies the presence of mostly B-particles sedimenting at 80S for the virus subject to control conditions (– PDS). In stark contrast, a substantial fraction of the virus treated with PDS shifted towards higher sedimentation rates with a peak in-between 150S (where the native virus sediments) and 80S, corresponding to (viral RNA containing) A-particles (hash). These results again point to a substantial impairment of in vivo RNA release by the incorporated PDS.
The observed effect of PDS in the PaSTRY assay and on the in vivo uncoating of RV-A2 was most likely due to the trapping/stabilisation of GQs located in the encapsidated RNA rather than an unspecific binding to other components of the virion. However, in one report, PDS was found to act as a weak inhibitor of the C5 convertase, a component of the complement system 79. Therefore, to further furnish our hypothesis, we repeated the in vivo analysis at capsid breathing conditions with another frequently used GQ-binding compound, Phen-DC3 80. RV-A2 was incubated as described above with PDS at 20 µM and 200 µM and in parallel with PhenDC3 at 1 µM and 5 µM, respectively. A mock-treated virus was used as a control. All samples were subjected to repeated centrifugal ultrafiltration to remove the excess of the compounds maximally. HeLa cells were then challenged with these samples and incubated for 9 h to allow for a one-cycle infection. RV-A2 positive cells were determined by fluorescence-activated cell sorting (FACS), using the intracellularly produced VP2 as a readout. Figure 3e demonstrates a concentration-dependent decrease in the number of cells containing replicating RV-A2 upon pretreatment with PDS. Notably, a significant reduction was also observed with 5 µM of Phen-DC3. This result further strengthened our hypothesis that PDS triggers a structural change in the encapsidated RNA as postulated from the PaSTRY analysis, preventing its orderly egress from the capsid under in vivo conditions.
PDS affects the conformation of the free RV genome and reduces viral infectivity in the presence of Na + but not in the presence of K+.
We next embarked on an ultrastructural analysis of gently extracted rhinoviral RNA for directly visualising the proposed PDS-induced structural change. The protein shell of RV-A2 was proteolytically removed with proteinase K. To additionally examine a possible differential impact of the prevalent extracellular (Na+) and intracellular (K+) monovalent cation, the digestion was performed with the purified virus in sodium- as well as in potassium-only containing phosphate buffer.
Initially, we evaluated the released naked (ex virion) viral RNA for the presence of GQs by mixing with ThT. Importantly, due to its moderate affinity, ThT binds only to already established GQs 56. At 30°C, this resulted in a significant increase in the fluorescent signal compared to the background (Fig. 4a, upper panel). However, the formation of some GQs in the viral RNA might be at least partially prevented by sequestration of the G-rich regions into kinetically trapped competing structures. In order to assess this possibility quantitatively, we heated the ex virion RNA for 10 min to 60°C (the temperature where it unwinds to escape from the RV-A2 capsid 73) in the presence of ThT followed by its slow refolding at room temperature, thereby promoting the formation of the thermodynamically most stable secondary structures. Under these conditions, the fluorescent emission intensity of the light-up probe substantially raised by about 20-fold in both samples (Fig. 4a, lower panel). This indicated that distinctly more GQs were now available for ThT binding, likely resulting from a conformational transition of kinetically trapped, metastable alternative (e.g. hairpin) structures to the more stable quadruplex structure triggered by the elevated temperature. A similar result was recently obtained in a study examining the kinetic vs thermodynamic control of a sequence within an mRNA able to switch from a hairpin to a GQ, using N-methyl mesoporphyrin (NMM) as light-up indicator 81. The recorded ThT signal was consistently higher for the Na+ compared to the K+-containing samples, despite the greater GQ stabilising propensity of the latter cation, amounting to a ~ 40 % difference at 30°C, which diminished to just 9 % for the 60°C heated samples. It suggests that the viral RNA molecule with intact 3D structure is possibly more compact in the presence of the charge-neutralising K+, leading to reduced accessibility of the already existing GQs for the ThT probe. Binding of ThT to GQs forming during refolding of the heated viral RNA was expectedly much less affected by the monovalent cation type when the probe was already present before full compaction of the nucleic acid molecule.
We next incubated the ex virion RNA with 20 µM PDS, again in a Na+ or K+-containing phosphate buffer, and submitted it to rotary shadowing. The platinum replicas were then observed by transmission electron microscopy (TEM) (Fig. 4b). In the absence of PDS, the ex virion RNA remained compact and approximately spherical though slightly deformed in Na+ (insets), indicating that compacting Mg2+ 82 and polyamines bound to the encapsidated genome 83, 84 were not removed by our extraction procedure. A PaSTRY analysis with these samples demonstrates the maintenance of a tertiary structure organisation under these conditions (see below). Strikingly and entirely in line with the proposed pyridostatin-induced RNA reorganisation deduced from the PaSTRy experiment (performed in PBS), PDS in the Na+-containing buffer led to substantial elongation of these RNA cores, making them appear irregular rods (left panel). Unexpectedly, no such effect of PDS was seen in the K+-containing buffer, where the RNA remained roughly spherical (right panel).
We then employed atomic force microscopy (AFM) imaging to independently confirm the TEM analysis outcome. AFM scans of identically prepared samples indeed yielded very similar images (Fig. 4c and Supplementary Fig. S7). They show mostly irregular rods in Na+-containing buffer + PDS with lengths of up to 200 nm and a diameter of about 10 nm (Supplementary Fig. S7, upper panels) and spherical RNA in K+-containing buffer + PDS with a diameter of roughly 25 nm (lower panels), compatible with the dimension of the capsid internal cavity. This confirms that the structures shown in Fig. 4b are specific for the viral RNA and not artefacts of the platinum contrasting and/or the drying process.
The drastic shape change only observed in Na+ containing buffer likely results from the rescuing of unstable GQs and/or the shifting of long-lived metastable structures with alternative G-quartet forming ability to the more stable GQ conformation by PDS. This presumably disturbs short and long-range interactions determining the global structure of the RNA genome 85, 86, similarly as described for certain small molecules on binding to tRNA and riboswitches 87, 88. Possibly, K+ binding pockets in the viral RNA as e.g. found in ribosomal RNA 89 further constrain and condense its native tertiary structure compared to Na+, limiting the access of the compound to these regions to account for the observed different effect of PDS. To substantiate this hypothesis, we assessed the stability of the protein-free (ex virion) RNA by carrying out a differential scanning fluorimetry (DSF) similarly as described by Silvers, Keller 90. Inspection of the DSF traces obtained in Na+ and K+ buffer indicated similarly low accessibility of the viral RNA for SYTO 82 at 25°C up to about 40°C, attesting that the RNA molecule stays compact and extensively folded within this temperature range (Supplementary Fig. S8). Here, SYTO 82 intercalates mostly into solvent-accessible double-stranded regions located at the periphery, which seems less affected by choice of these cations compared to the accessibility of GQs for ThT under similar conditions (Fig. 4). The subsequent relatively rapid fluorescence increase from the lower baseline (at 44.5°C and 51°C for Na+ and K+, respectively) to the upper base line (= maximal response; at 54.3°C and 57.1°C for Na+ and K+, respectively) relates to the disruption of (mostly) the RNA tertiary structure, now allowing SYTO 82 binding also to internally localised stem regions. The rise was ~ 2-fold (Na+) and ~ 1.2-fold (K+), indicating a more extensive unfolding in the sodium-containing buffer. The higher stability of the tertiary contacts of the viral genome in the presence of K+ is illustrated by the around 4°C higher Tm1. The following progressive drop in emission intensity is due to melting of the more stable secondary structures with the release of SYTO 82, resulting in somewhat closer spaced higher melting temperatures Tm2 (60.5°C for Na+ and 63°C for K+) for this transition.
The above in vitro data derived with the ex virion RNA altogether implies that its distinctly more compact tertiary structure imposed by K+ hinders the PDS molecules from arriving at internal regions able to transform into GQs on the binding of this compound. If correct, this should materialise in a distinctly smaller number of PDS molecules associated with the RV-A2 genome in potassium compared to sodium buffer. As assessing this figure is technically less demanding with encapsidated RNA, we quantified the amount of PDS that remained associated with the virions after incubation (20 µM) at 4°C or at 34°C for 4 h in Na+ or K+ containing buffers and subsequent extensive washing to remove any externally deposited traces of the compound. Table 3 shows that the number of PDS molecules, which penetrated the capsid upon incubation at 34°C in Na+-containing buffer was by far higher than in all the other incubation conditions.
Table 3
– Mass spectrometry quantification of PDS content per virion.
Treatment
|
PDS bound (moles/mole virus)
|
Relative to maximum
|
34°C / Na+
|
10
|
100 %
|
4°C / Na+
|
0.9
|
8.8 %
|
34°C / K+
|
0.2
|
1.4 %
|
4°C / K+
|
0.4
|
3.7 %
|
The result further corroborated our hypothesis that K+, while commonly cooperating with PDS in GQ stabilisation 91, unexpectedly protects the rhinoviral RNA from binding of this compound not only ex virion but also inside authentic viral particles (in virion). The magnitude of the difference of virion-incorporated PDS was quite impressive, as the encased polynucleotide of picornaviruses is already very compact 92, 93. This might have conceivably mitigated the tertiary structure changes in the presence of Na+ vs K+, believed to govern the extent of pyridostatin binding to viral QGRS when free in solution. A monovalent cation-dependent breathing activity as an additional source for that difference was ruled out by a nanoDSF analysis of purified virions diluted in 100 mM sodium or potassium phosphate buffer (Supplementary Fig. S9).
We then assessed whether the different PDS uptake by RV-A2 as found by MS resulted in consequences on its infectivity by determining the TCID50 of virus exposed to the compound diluted in the respective monovalent cation-containing phosphate buffer. To avoid any unspecific loss of infectivity due to thermal inactivation, we reduced the incubation temperature to 25°C and compensated for the resulting diminished breathing by extending the treatment time to 20 h. As shown in Fig. 4d, the virus sample treated with PDS in the presence of Na+ exhibited a titer reduction by 2 logs compared to control conditions (the same buffer without PDS). By contrast, PDS treatment in K+ buffer did not affect the virus titer as expected from the small amount of the compound detected by MS in the virion of similarly treated virus (Table 3). Note that the slight reduction of infectivity on changing the internal monovalent cation environment from Na+ to K+ in the absence of the PDS was not significant.
Finally, we explored at what step of the infection cycle PDS might act on RV-A2 in vivo (i.e. without pre-incubation) by a time-of-addition experiment. The virus was bound to the cells for 30 min at 4°C, PDS was added, and the cells shifted to 34°C (T0). The same experiment was conducted in parallel, except that PDS was added at T180 and T300, respectively. This roughly corresponds to RV entry and uncoating (T0), RNA synthesis (T180), and assembly (T300) 94, 95, 96. The cells were maintained for 9 h pi (one full cycle of infection), and viral synthesis was measured by fluorescence-activated cell sorting (FACS). As can be seen in Fig. 4e, the infection rate and viral synthesis was only significantly impacted upon the addition of PDS at T0.